Chapter 41Markers of Infection

Graham DY, Qureshi WA.

Publication Details

Markers for the presence of a Helicobacter pylori infection consist of features or events that suggest that the infection is, or has been, present and include alterations in gastroduodenal structure and function, as well as tests to detect the bacterium itself or the presence of bacterial enzymes. Initially, the primary marker suggesting an H. pylori infection was the presence of one of the diseases associated with the infection, including histologic gastritis, peptic ulcer disease, and gastric adenocarcinoma. It is now recognized that primary mucosa-associated gastric lymphoma (MALT lymphoma) and approximately 10% of cases of nonulcer dyspepsia are also directly related to H. pylori infection. Once it was confirmed that H. pylori was not a colonizer of inflamed gastric mucosa, but was rather the cause of gastric inflammation (gastritis), it was only a matter of time before the previous associations with gastritis could be transferred to H. pylori and prior associations clarified.

Laboratory Abnormalities

Serum Gastrin or Pepsinogen

H. pylori infection is associated with progressive and complex abnormalities of gastroduodenal physiology. There are a number of biochemical markers suggestive of the presence of an active or past H. pylori infection. For example, one biochemical marker of an active H. pylori infection is an increase in fasting and in meal-stimulated serum gastrin levels (40, 63, 64, 90, 100). Prior to the discovery of H. pylori, elevated pepsinogen I levels were thought to be a genetic marker for risk of developing duodenal ulcer disease (94). Clarification of the role of H. pylori-associated gastritis in gastrin release showed that an elevated serum pepsinogen was not a genetic marker relating to duodenal ulcer but rather was related to the presence of the infection (4). Intrafamilial clustering of H. pylori infection led to the erroneous conclusion that it was a genetic marker. Marked or extreme hypergastrinemia is associated with destruction of the stomach, leading to profound hypochlorhydria, and can be seen in long-standing infection with H. pylori, but it is not specific because it is also present in autoimmune gastritis (99).

Several investigators have suggested that measuring the change in gastrin or pepsinogen levels following treatment might be a useful method to evaluate whether the infection had been cured (12, 15, 16, 38, 40, 41, 66, 79). For example, Furuta et al. (35) suggested using a 25% increase in pepsinogen I/II ratio as the cutoff for determining whether the infection had been cured. They reported a sensitivity of 95.7% and a specificity of 89.7%. This hypothesis was tested recently using paired sera to evaluate the changes in gastrin levels, as well as in serum pepsinogen I and II levels and pepsinogen I/II ratios and IgG titers after successful treatment (2). The decline in meal-stimulated gastrin was significant 2 weeks after therapy. Cure of infection also resulted in a significant fall in levels of both fasting and postprandial pepsinogen I and II levels. The fall in pepsinogen I/II ratio was the most marked change. Nevertheless, none of the parameters proved useful to detect cure of the infection in an individual patient because, despite statistically significant changes in pepsinogen and gastrin levels for the group as a whole, no cutoff value or percent change was found that reliably identified whether the infection had been cured (2). Measurement of a change in levels of pepsinogen or gastrin requires the use of paired sera and waiting for more than 6 months (likely more than 1 year) to obtain reasonably accurate results, and none of these tests can be recommended for clinical use, either diagnostically or to test whether the infection has been cured. There are a number of noninvasive tests that can provide an accurate assessment of H. pylori status pre- and posttherapy (see below).

Inflammation

H. pylori causes chronic inflammatory destruction of the stomach and, as one would expect, there are markers of this inflammatory process (88). Some are subtle and can be found by comparison with control groups of uninfected individuals or occur after H. pylori eradication. For example, several studies have suggested that there is a reversible fall in leukocyte count after cure of the infection (42, 51). In a recent study looking at the effect of H. pylori infection and CagA status on leukocyte counts and liver enzymes, pretreatment white blood cell counts and serum aspartate transaminase (AST) were higher in infected patients (42). The AST levels were higher in infections with CagA-positive strains compared to CagA-negative strains. When treatment resulted in a cure, there was a significant fall in the total white blood cell count from 7,413 per mm3 to 6,738 per mm3 (P = 0.04). This drop was accounted for largely by a reduction in the polymorphonuclear leukocyte count. The AST levels were unaffected. It is important to note that the white blood cell count and the AST levels were within the normal range before and after therapy, suggesting that the "normal" range may reflect use of many infected individuals. The fact that the AST did not change suggests that it was actually not related to the infection or that it is a marker for susceptibility to infection with CagA-positive H. pylori.

Effect on Acid Secretion and Histology

H. pylori infection damages the gastric mucosa and has an effect on gastric acid secretion. The overall effect is related to the predominant form of gastritis. The stomach can be divided conveniently into an acid-secreting portion (the body or corpus) and the distal stomach where acid-secreting cells are either absent or sparse (the antrum). H. pylori organisms are present over the entire surface of the stomach, but the interaction of the bacterium and the surface cells differs among the regions. The severity of damage in the antrum is always equal to and typically more severe than that in the corpus (36). High levels of acid secretion are associated with mild corpus inflammation and low H. pylori density in the corpus. This is typical of patients with duodenal ulcer disease and has been termed the antral predominant or corpus-sparing pattern of gastritis. Inhibition of acid secretion allows the H. pylori to interact with the mucosa, producing an increase in corpus inflammation irrespective of the method used to reduce acid secretion.

Typically, in H. pylori infection there are reversible abnormalities in acid secretion, with acid secretion being prolonged after meals due to a defect in the reflex inhibition of acid secretion as the intragastric pH falls below 3 (19). Eradication of the infection restores the normal inhibitory pathways and may have other effects on acid secretion depending on the pattern of gastritis. For example, inflammation of the gastric corpus inhibits acid secretion, and following cure of the infection, the acid secretion produced by a stimulus such as pentagastrin increases markedly (44). In contrast, when the corpus is minimally inflamed, there is typically either no change or a slight decrease in pentagastrin-stimulated acid over time, possibly related to the resolution of the increase in gastrin secretion (19). It is thought that the inflammation-reduced acid secretion from the corpus is related to production of the cytokine interleukin 1 (IL-1).

Histologic Markers of the Infection

Histological gastritis is the primary manifestation of the infection. The typical histology is infiltration with a combination of acute and chronic inflammatory cells along with development of intramucosal lymphoid aggregates and follicles (88). H. pylori cells are present in sufficient quantity to be seen with high-power or oil-immersion magnification of histologic sections or of Gram stains of smears of gastric mucus. The typical histologic stain, hematoxylin and eosin (H&E), is excellent for identifying the presence of gastritis, and the clinical sequelae of the infection such as gastric carcinoma or MALT lymphoma, but is rather unreliable for identifying the actual bacterium. Special stains to identify H. pylori are preferred to reduce error. Special stains are especially helpful posttherapy when there is still considerable remaining chronic inflammation. The Warthin-Starry and modified Giemsa stains are widely used to make identification of the bacteria easier, but neither is ideal. Triple stains which combine H&E, Alcian blue, and another stain to identify the bacterium such as the Genta or El-Zimaity triple stains allow easy identification of H. pylori and excellent visualization of gastric morphology (13, 24, 25, 27, 37). The original Genta stain used uranyl nitrate, which is not available widely, and the staining technique was not suitable for use with an autostainer. Recent modifications include replacing uranyl nitrate with a lead nitrate-gum mastic solution and using a microwave oven for the sensitization, silver impregnation, and reduction steps. This reduced the staining time to 28 min. The technical time can be reduced to 9 to 10 min if deparaffinzation and all steps following reduction are done with the help of an autostainer (27).

Our laboratory routinely uses the Genta or El-Zimaity triple stains, which both require only one slide. For those laboratories that use a special stain only when there is a possibility of H. pylori (e.g., when the mucosa shows inflammation) and posttherapy, we recommend the use of the Diff-Quik stain as it is readily available, rapid, and inexpensive (25). When only a few bacteria are present and there is question about whether a case is positive or negative, we use the El-Zimaity dual stain, which combines periodic acid-Schiff (PAS) and a silver stain (26) (Fig. 1). This is the only practical way to visualize H. pylori in the duodenal mucosa. Failure to include a special stain may cause a false-negative result in up to 25%, especially in posttreatment patients. Immunohistochemistry is used in some laboratories, but it is inferior to the triple stains or the dual stain for H. pylori diagnosis.

Figure 1. Example of the El-Zimaity dual stain of gastric mucosa (26).

Figure 1

Example of the El-Zimaity dual stain of gastric mucosa (26). The stain combines periodic acid-Schiff and a silver stain and is the preferred stain when the density of H. pylori is very low or when searching for H. pylori in the duodenum.

A group of expert pathologists recommended that for diagnosis of H. pylori infection and characterization of the pattern of gastritis, two biopsies should routinely be taken from the antrum and two from the body of the stomach for histological evaluation (18). These biopsies should be taken from the most normal-appearing mucosa. For detection of H. pylori, we routinely take three biopsies, one each from the distal antrum, the angulus incisura, and the greater curve of the mid corpus as this has an essentially 100% accuracy for detection of H. pylori infection (22, 23). Whenever possible, large-cup biopsy forceps should be used. Since the organisms are most abundant in the mucous layer on the surface of the tissue, it is important not to wash off this layer. We recommend that biopsy specimens not be handled or oriented by the endoscopist but rather be "shaken off" the forceps by shaking the opened forceps in formalin (Table 1). The biopsy samples are fixed immediately. It is important to train the technicians to mount the specimens "on edge" so that the surface can be visualized. Several cuts are examined. False-negative results may occur due to incorrect sample collection, such as biopsy from an area of atrophy or metaplasia, or recent treatment with proton pump inhibitors or antibiotics. Taking specimens from the gastric corpus, which is less frequently involved by intestinal metaplasia, is especially important in geographic regions where atrophic gastritis is common.

Table 1. Recommendations for collecting gastric mucosal biopsies for detection of H. pylori.

Table 1

Recommendations for collecting gastric mucosal biopsies for detection of H. pylori.

Markers Based on H. pylori Enzymes

Rapid Urease Tests

H. pylori contains abundant urease, making the presence of urease activity a useful marker for the presence of the organism. When a biopsy specimen containing H. pylori is introduced into a medium containing urea, urease splits the urea into ammonia and carbon dioxide. The ammonia released results in an increase in pH, which can be detected by a color change of a pH indicator (Fig. 2). The accuracy of rapid urease testing is high, such that a correlation between histology and rapid urease testing provides a simple measure to gauge the accuracy of the pathology department. In a study with 143 patients, we found the sensitivity and specificity of different rapid urease tests to be in the range of 95 and 100%, respectively (22). There is little difference among the available rapid urease tests, so that cost and availability are the prime determinants of which is used. The speed of the reaction is enhanced when large biopsies or multiple biopsies are placed in a single test well and when a warmer is used to speed the reaction (61, 62, 113, 114). The sites with the highest yield are the gastric angle and the greater curve of the corpus (109). One approach is to place a sample from each of these sites into the same well of the rapid urease test.

Figure 2. Examples of three commercial rapid urease tests including the original CLO test and two second-generation tests.

Figure 2

Examples of three commercial rapid urease tests including the original CLO test and two second-generation tests. One of each pair is the control, the other is after the color change.

Rapid urease tests require a high bacterial density such that recent use of antibiotics, bismuth-containing compounds, or proton pump inhibitors may cause false-negative results. Because of the high cost of histologic examination in the United States, it has been suggested that it may be cost effective to retain the biopsy specimens for histology in the endoscopy laboratory until after the results of the rapid urease test are known. If the rapid urease test is negative after 24 h, the biopsy specimens are sent to the laboratory. If the rapid urease test is positive, they are discarded. Of course, biopsy samples from abnormal-appearing mucosa must always be sent to the laboratory for processing and examination.

Urea Breath Testing

The urea breath test is the noninvasive method of choice to determine H. pylori status either pre- or posttherapy. This test is based on the organism's urease activity, which liberates CO2 from labeled urea, resulting in the production of labeled CO2 that can be easily detected in the breath. Two forms of labeled urea are commercially available: one contains the stable, nonradioactive isotope 13C and the other contains the radioactive isotope 14C. Although the amount of radiation exposure with the [14C]urea breath test (14C-UBT) is small, none is best and the test is contraindicated in children, as well as pregnant women and, possibly, women of childbearing age. The amount of radiation exposure is approximately equivalent to one day's background radiation, but as the label can be potentially incorporated into the bicarbonate pool and the half-life is very long, the long-term effects are unpredictable, making the 13C test preferred, where available.

The 13C-UBT has proven to be an extremely reliable test and yields satisfactory results despite almost every conceivable modification that has been tried. The urea breath test is a qualitative, not a quantitative, assay for the presence of gastric urease activity. The concepts underlying the test are straightforward, as are the factors that could potentially lead to false-positive or false-negative results (Table 2). Urease activity is not present in mammalian tissue but is widely distributed among bacteria (e.g., the microbiota of the mouth contain many urease-containing bacteria). A number of factors could theoretically affect the 13C-UBT results (Table 2), including exposure of urea to oral microbiota, either in the mouth or the stomach, which could hydrolyze the substrate and cause a false-positive urea breath test result. To overcome this problem, some investigators have encapsulated the urea or administered it as a tablet (6, 45). These routes of administration have, in turn, raised other issues of concern related to the dissolution and distribution of tablets and capsules within the stomach as well as the possibility of emptying prior to dissolution, leading to false-negative results (43). Other investigators have instructed patients to cleanse their mouths before ingesting the urea to reduce the number of bacteria, but this procedure would not eliminate exposure to those organisms already ingested (77, 85). The pH optimum for the urease of non-H. pylori gastric contaminants is generally 7 or above, and urea hydrolysis will not occur in the stomach unless the quantity of bacteria and the pH are both high. The potential impact of mouth bacteria can be overcome by delaying the first post-urea breath sample for 15 or 30 min in order to dilute any swallowed bacteria and to allow the acid in the stomach to inactivate mouth bacterial ureases. Administering the urea after a test meal provides similar benefits since the meal causes dilution of the gastric contents and secretion of acid, which lowers the pH. Despite these precautions, both the quantities of bacteria and the pH may be high in patients with gastric atrophy and result in false-positive urea breath tests associated with negative serologic results (80).

Table 2. Considerations regarding the accuracy of the 13C-urea breath test.

Table 2

Considerations regarding the accuracy of the 13C-urea breath test.

The specificity and sensitivity of the standard U.S. protocol for the 13C-UBT have repeatedly proved to be excellent, providing reliable information about H. pylori status before or after therapy (55, 56, 58). Recent studies have shown that it is possible to retain the accuracy of the 13C-UBT while simplifying the test. Simplifications included elimination of a special breath collection bag and collection of breath samples using a straw to blow into a test tube, elimination of fasting for more than 1 h before testing, and elimination of the solid test meal. These changes have had the practical advantage of allowing a shorter test period as well as enhancing the convenience to the subject and the person administering the test. Replacement of the solid test meal with a citric acid solution eliminated meal preparation and results in a solution with a pH below 4 to inhibit ureases other than H. pylori and to stimulate urease activity, probably by increasing the permeability of the membrane through the postulated Urel channel (29, 78, 96).

The amount of substrate used in the 13C-UBT has not proven to be a critical factor for achieving accurate results. Theoretically, there is a lower limit below which the proportion of false-negative tests would increase, but varying the amount of expensive [13C]urea from 250 to 75 mg has been done without a reduction in either specificity or sensitivity. This mass of substrate administered ensures that substrate exhaustion does not occur such that breath sampling can be delayed until any pulse of labeled CO2 from orally hydrolyzed urea has passed (58, 69). Extra or "cold," or nonradioactive, urea was not added to the commercial 14C-UBT, which is administered as a capsule containing carrier-free radioactivity. With that test even a small quality of urease can cause substrate exhaustion.

Breath collection devices

A number of different collection devices have been used to collect breath samples for analysis. The type of breath collection device used is dictated by the requirements of the analyzer used and the need for storage or shipment. Overall, collection devices have not proven to be a problem. Typically, when mass spectrometry is used for analysis, the breath samples are shipped to a central facility in glass or plastic tubes similar to those used for blood collection. The original breath sample is either collected in a bag and a small sample is transferred to the shipping tube or the breath is collected directly in the tube by blowing through a straw inserted into the bottom of the tube, which is then capped (7). When larger breath samples are required, as in the use of infrared instruments, a CO2-impermeable bag of 150 to 400 ml is used.

Detection of 13CO2

As previously described, the 13C-UBT detects H. pylori infection indirectly based on the hydrolysis of orally administered [13C]urea. The liberated CO2 appears in expired breath where it can be detected as an increase in the isotopic ratio of 13CO2/12CO2. Thus, in the 13C-UBT a breath sample is taken before the substrate is administered and another sample is taken 15 or 30 min after, depending on the test kit. Classically the enrichment of 13CO2 in the breath has been assessed using a gas isotope ratio mass spectrometer. Such instruments require very little sample and are extremely sensitive and accurate even at very low levels of respiratory CO2. Newer methods based on infrared spectrometry also can be used to determine the ratio of 13CO2/12CO2 (9, 49, 84, 86). These offer the potential of lower instrumental costs and for analysis at the point of patient care. The choice of analyzer depends on the number of samples to be analyzed as well as the characteristics of the analyzer. Many of the infrared instruments are capable of detecting the enrichment of 13CO2 in the breath when the amount is well above the cutoff value but are less able to provide an accurate estimate of samples near this value. In Western countries, evaluation of dyspeptic patients for the presence of H. pylori is likely to become one of the most frequent indications for requesting a urea breath test. In this population the frequency of H. pylori is typically low (e.g., the proportion of white United States–born individuals is less than 12%). Because the majority of test results will be negative, the test must be accurate in the region around the cutoff value. This region is precisely where mass spectrometry excels and some of the newer instruments have the greatest difficulty. In clinical trials this has not proven to be a significant problem; nevertheless, it behooves the physician who contemplates purchase of a point-of-care instrument to ensure that it provides acceptable accuracy in the range around the cutoff point. One approach to work around this problem is to send samples with results near the cutoff value for reanalysis by mass spectrometry.

Expression of results

In the United States, the cutoff value for the 13C-UBT in adults was determined using receiver operation characteristics (ROC) curves to determine the lowest increase in 13CO2 abundance associated with H. pylori infection in a group of 60 H. pylori-infected and 60 uninfected volunteers (58). The cutoff for a positive test was defined as an increase of >2.4%, and this was validated against histologic examination and culture of gastric mucosal biopsies (58). Nevertheless, calculation of the results in terms of an increase in 13CO2 abundance may not be the best method. In other 13C-UBT applications it has been customary to calculate the proportion of the dose recovered in breath. This calculation takes into account the weight of substrate, its molecular weight, number of labeled carbons, the rate of CO2 production, as well as the breath enrichment of 13CO2 at a given interval after substrate ingestion. Although the simple change in isotopic abundance from the pretest baseline value to postdose value initially sufficed for diagnostic purposes in adults, it poses problems particularly in the use of the test in children. The test outcome hinges on two separate processes, one arising from the organism and the other from the host. H. pylori urease liberates pure 13CO2 from the administered [13C]urea. As the labeled CO2 combines with respiratory CO2 at its natural abundance, the degree to which the 13CO2 is diluted by host CO2 will determine the degree of enrichment present in the expired breath. CO2 production differs in relation to age (adults greater than children), sex (males more than females), weight, or height. Thus, the identical amount of [13C]urea hydrolyzed in the stomach of a small person (e.g., child) would provide a proportionally higher enrichment than that from a large person. Any cutoff value based on breath enrichment alone may give rise to misleading results when used in populations whose anthropometric measurements differ significantly from those of normal U.S. adults. This problem can be overcome by using established methods to estimate an individual's CO2 during a breath test. One method is based on body surface area (in m2) and is calculated from measurements of height and weight. The method assigns a value of 300 mmol CO2/min/m2 (48). The second uses equations developed by Schofield to predict basal metabolic rates from the age, sex, height, and weight and has the advantage of six age stratifications (95). With the Schofield equations the rate of actual urea hydrolysis can be obtained by combining the estimate of host CO2 production rates with urea dose and breath enrichment values. When this is done, the cutoff value is expressed as the urea hydrolysis rate (μg/min), and a value of 10 μg/min provides indication of the presence or absence of H. pylori infection that is independent of age, sex, height, or weight (57). Use of this test result expression is recommended and is absolutely essential for children under the age of 5. One cannot use the absolute level of urease activity to predict the number of H. pylori in the stomach in more than the most general way.

Use of the urea breath test

The urea breath test, histology, and rapid urease testing all require a relatively high density of H. pylori. Thus, any practice that reduces the intragastric concentration of H. pylori could lead to false-negative results, and avoidance of low concentrations of H. pylori is the basis for the recommendation to withhold confirmation testing until 4 or more weeks after ending therapy. Use of bismuth or antibiotics in the pre-testing period may also result in a false-negative test. Proton pump inhibitor therapy reduces the density of H. pylori through a direct anti-H. pylori effect and approximately 20% of H. pylori-infected patients will have a false-negative result when taking a proton pump inhibitor. We recommend that proton pump inhibitor therapy be stopped at least 1 week before testing. In our experience and the experience of others, standard dose H2-receptor antagonists do not affect the accuracy of the 13C-UBT and they can be used up to the time of testing (10). False-positive tests are rare, but when they occur, they are likely due to contamination of the stomach with urease-positive organisms. This can be reduced by using the citric acid test meal because, as noted above, this reduces the intragastric pH below the pH optimum of these ureases.

Markers Based on Anti-H. pylori Antibodies

Serologic Markers

There have been a number of methods developed for the noninvasive detection of anti-H. pylori IgG or IgA, including whole blood, serum, saliva, and urine (17, 30, 33, 98, 107), as well as immunoblotting (5, 11, 32, 52, 54, 82, 92, 101). Enzyme-linked immunosorbent assays (ELISA) for urine IgG have also proved successful but have the same drawback as serum methods (3, 53, 76). Detection of anti-H. pylori antibody is the easiest noninvasive approach to test for the presence of an H. pylori infection. With some tests only a few drops of fingerstick blood are required and the results are available in less than 5 min.

Antigens

The initial serologic tests were patterned after those developed for Campylobacter jejuni and consisted of a mixture of very crude antigens. Cross-reactivity with other bacterial antigens (e.g., flagellar antigens) was expected and indeed occurred. Second-generation tests have used purified antigens such as the high-molecular-weight antigens, which include antigens that are confirmationally determined. Some of the second-generation tests have proven accurate worldwide (28, 74, 102, 106, 108). ELISA are widely available because of their low cost, rapidity, and reproducibility. Complement fixation, hemagglutination, bacterial agglutination, immunofluorescence, and immunotransference (Western blot) tests are also available but are less accurate and are not widely used. Tests using crude antigens (e.g., "in-house" tests) are generally less reliable and are not recommended for clinical use. Tests for specific virulence factors such as CagA have been developed but have not proven to have any special clinical utility (47) and have generally suffered from poor specificity and sensitivity (111, 112).

Interpretation of results

Detection of serum IgG against H. pylori typically indicates a current or prior infection. Recent studies have shown that the prevalence of H. pylori infection is decreasing in all age groups in both Japan and the United States (60, 70–72) and likely in all Western countries. Thus, the population of individuals with anti-H. pylori antibodies and negative UBTs or no H. pylori seen on histology has increased and will continue to cause an underestimation of the true ability of serologic tests to accurately identify the presence of anti-H. pylori antibodies in body fluids. In our experience, those with positive ELISA using second-generation ELISAs, such as HM-CAP, typically have histological evidence of an old H. pylori infection (e.g., lymphoid aggregates). When serologic tests that are accurate in Western countries are used in other countries such as in Japan, the cutoff value must often be adjusted, usually upward. We speculate that this is not because the antigens of different H. pylori infections differ remarkably, rather, the cutoff must be raised to exclude those whose infections have been lost and whose antibody titers are falling. The accuracy of any test is only as good as the tests used to confirm the correctness of the serologic test. For example, in Japan the results of serologic tests were markedly influenced by the histological techniques used to determine the presence or absence of active H. pylori infection (75). It is now apparent that it is insufficient to only be able to accurately diagnose the presence of an active H. pylori infection (e.g., histologically or with UBT); to characterize a serologic test one must also take into account evidence of prior H. pylori infections. One can therefore never expect serologic tests to provide as accurate results for the presence of an active H. pylori infection as do histology, culture, or the UBT. In addition, in general as the prevalence of the infection falls in a community, the accuracy of serologic tests suffers with an increase in the proportion of false-positive tests. In that population, a negative test essentially excludes an H. pylori infection (65).

Changes in titer following cure of the infection

Antibody titers fall following successful eradication of infection. A number of authors have investigated whether this can be used to identify whether an individual patient has been cured. Again, as with pepsinogen and gastrin levels, it is easy to demonstrate a fall in titer for the group under investigation but not for an individual patient (2). Paired sera and months of follow-up are needed, making this a clinically impractical approach. The decrease in titer after cure is slow and unpredictable, but over time most individuals will serorevert. In our experience this may require decades. For example, Al-Assi et al. found that although there was a drop in anti-H. pylori IgG antibody titer during treatment, levels in only one patient (6%) dropped below 50% (2). Kosunen et al., on the other hand, using the 50% drop in titer as an indicator of treatment success 6 months posttreatment, reported a sensitivity of 97% and a specificity of 95% (59).

Test for IgA and IgM Anti-H. pylori Antibodies

Although tests for IgA and IgM to H. pylori are available in many countries, none are approved by the U.S. Food and Drug Administration. Overall, neither IgM nor IgA tests alone are superior to IgG serology, and the sensitivity and specificity of these tests have generally been too low for them to be recommended either alone or in combination with an IgG test.

Markers Based on the Presence of H. pylori Antigens

Stool Antigen Testing

Recently there has been increased interest in identifying H. pylori protein antigens in stool as a marker of infection. Premier Platinum HpSA (Meridian Diagnostics Inc., Cincinnati, Ohio) has developed an in vitro qualitative enzyme immunoassay commercial kit that is stated to be able to detect H. pylori protein antigens of concentration ≥184 ng/ml of feces. The test uses polyclonal antibodies to H. pylori as capture antibodies, adsorbed to microwells. Samples are diluted as per manufacturer's instructions and added to each antibody-coated microwell. After incubation for 1 h with a peroxidase-conjugated polyclonal antibody, unbound material is washed off, substrate is added, and the wells are incubated for 10 min at room temperature. Color develops in the presence of bound enzyme that is measured spectrophotometrically after adding a stop solution. Absorbance is measured at 450/630 nm.

Overall, studies using pretreatment stool H. pylori antigen tests have shown that the sensitivity and specificity of stool antigen testing were comparable to histology or UBT (1, 8, 14, 31, 50, 81, 83, 87, 103–105). For example, in one prospective multicenter trial with 501 patients with H. pylori infection, proven by histology and a rapid urease test or culture, stool antigen for H. pylori was positive in 256 of 272 patients (sensitivity, 94.1%; 95% confidence interval, 91 to 97%). Of 219 patients without infection, 201 were negative by HpSA (specificity, 91.8%; 95% confidence interval, 87 to 95%). In that study the post-treatment sensitivity and specificity for this marker were lower (90 and 95%, respectively) (104).

There are exceptions reporting less satisfactory results (34, 73, 91, 93), and the optimum cutoff values for optical density have varied slightly between studies (34, 83, 87). It has now become evident that there may be considerable lot-to-lot variation in stool antigen tests. The most likely explanation is the polyclonal sera used for the capture antibody are obtained from rabbits and are thus difficult if not impossible to standardize. Stool antigen testing has proved to be less reliable when used soon after the end of therapy, and it is now generally recommended that one must wait longer to confirm eradication (39, 68). Generally, posttherapy studies have generally also shown good sensitivity and specificity when testing is delayed at least 4 weeks. The concept of a stool antigen test is a good one, and several companies have stool antigen tests in trial that use monoclonal antibodies as capture antibodies.

In general, one can use a UBT or the stool antigen for initial diagnosis, or as confirmation of a positive serologic result with the caveat that one should probably wait 6 or 8 weeks after therapy when using the stool antigen test. The UBT is preferred where available.

Markers Based on the Presence of the Bacterium

Culture

H. pylori is fastidious both in its growth and transport requirements. Successful culture relies on the transport medium, time in transit to the pathology laboratory, temperature during transportation, and the medium used; all influence bacterial viability and recovery (46, 110). For transportation and storage, various media have been evaluated. We use cysteine-Albimi broth with 20% glycerol and have found that skim milk with 17% glycerol was equally satisfactory (46). At room temperature there is a decrease in H. pylori titer after 6 h. At 4°C the organism will survive for 1 week and at −70°C, or in liquid nitrogen, it will survive indefinitely. Once plated on two media (selective and nonselective), the samples are incubated at 37°C with high humidity in a microaerophilic atmosphere for at least 10 or preferably 14 days. The first colonies appear after 3 to 4 days. Identification is by colony morphology, Gram stain, and the enzymes the bacteria produce, including urease, oxidase, catalase, and glutamyltranspeptidase. Culturing the organism also allows testing for antibiotic susceptibility. This is particularly useful in treatment failures or in areas of high antibiotic resistance. The specificity from culture is 100%. The sensitivity varies from 50 to 99% depending on the laboratory and interest of the microbiologist. Most laboratories with an interest have extremely high yields.

The order in which biopsies are taken (culture or histology) does not make a difference as preimmersion of biopsy forceps in formalin had no detrimental effect on the ability to culture H. pylori (115). Culture from stool has proven to be difficult, with a low yield (89). The frequency of positive culture might be enhanced by induced diarrhea (89).

PCR Tests for Markers of H. pylori Infection

Bacterial DNA can also be used as a marker for the infection. Under the ideal circumstances, the sensitivity is close to that of culture. The potential advantages of PCR include high specificity, quick results, and the ability to type bacteria without the requirement for special transport conditions. The routine use of PCR to diagnose H. pylori infection has proven problematic. PCR is very sensitive to inhibition by factors present in stool (20, 21). False-positive results are common, possibly because of contamination in the laboratory. Because PCR assays detect homologous sequences, if other bacteria contain those sequences, it will test positive, causing specificity to be low. Although this test has high sensitivity and is sometimes compared to the UBT, practical considerations and cost have limited its use (20). However, PCR use in H. pylori infection remains very promising and may permit rapid detection of antimicrobial resistance by detecting mutations associated with various antibiotics (67, 97). At the present time PCR remains a research tool for the diagnosis of H. pylori infection.

References

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